Why collect/salvage? — Herpetological collections are the basis for the study of herpetofaunal biodiversity. Each specimen is a verifiable permanent record from a particular place at a specific time. Collections facilitate the acquisition of knowledge in biogeography, systematics, evolution, and ecology. The fundamental role of herpetological collections is to safeguard the specimens they contain.

• Collections are curated repositories lasting in perpetuity.
• Specimens are references that are shared, loaned to, and exchanged with other scientists.
• Collections constitute a baseline, without which current comparisons could not be made nor predictive models tested.
• Modern collections incorporate genetic resources for their specimens.
• Series of specimens allow herpetologists to study morphological and genetic variation and correlate the two.
• Herpetological collections do not negatively affect amphibian and reptile populations.
• Herpetological collections provide the data to conserve amphibian and reptile populations.

Scientific specimens are only as good as their associated data and field notes. Keep them organized, and don't let your valuable specimens become teaching collection objects.

Permits — To legally collect/salvage/possess native Kansas amphibians and reptiles for non-commercial purposes, you need to have the proper permits/licenses. You will also neeed to adhere to the bag and possession limits until the specimen is transferred to a museum.

• State-listed species (Endangered, Threatened, or Species in Need of Conservation)
  • To possess state listed herps you will need to apply for and obtain a Scientific Collecting Permit from KDWPT.
• Non-native herp species
  • There are no restrictions on collecting/salvaging non-native herps.
• Other native Kansas herp species
• Kansas Residents, 16 years old or greater
  • American Bullfrog (Lithobates catesbeianus)
    • A fishing license in season (1 July through 31 October).Daily creel limit 8 frogs; possession limit 24 frogs.
  • Snapping Turtle (Chelydra serpentina), Smooth Softshell (Apalone mutica), Spiny Softshell (Apalone spinifera)
    • A fishing license in season (currently year-round). Daily creel limit 8 of any combination; possession limit 24 in any combination.
  • All other native non-state listed Kansas herps
    • Hunting license in season (currently year-round). Possession limit 5 per species.
• Kansas Residents, under 16 years old
  • Subject to the same daily and possession limits, but do not need to purchase/possess licenses.
• Non-Residents
  • Subject to the same daily and possession limits, but need to purchase/possess non-resident licenses.

What to collect — All specimens are valuable but some have more inherent (or immediate) value than others. In general, the more 'rare' a specimen is perceived to be, or the greater it extends the known range or fills a gap... the more value it is scientifically. Additionally, it is important to collect/salvage specimens before they are even needed, so they may be available when requested later. This is both an aid to the eventual researcher but also helps ensure that Kansas is well-represented in any future analyses. It is seldom practical (or necessary) to collect/salvage every specimen. One should consult the range map in the KHA and weigh the significance of the new specimen in helping to better define the species range in Kansas.
Condition is also an issue: road-killed snakes that have been hit just once make great specimens... while turtles and amphibians less so. A high priority specimen (based on location and perceived rarity) should be salvaged regardless of condition.

Euthanasia — Live animals will need to be euthanized before their preparation as scientific specimens. Many chemicals may be used for anesthesia (inhalation and injection), however, whole-body cooling, followed by freezing remains one of the most humane and practical methods. Whole-body cooling/freezing is a pain-free method to dispatch herp specimens (Lillywhite, et al. 2017. [Bioscience. 67(1): 53-61 ]).
Care should be taken to avoid freezer burn (dehydration). Place specimens in an air-tight plastic bag. Specimens can be kept this way for more than a year without issue. Cloth bags should be avoided, as the specimens will dry out quickly and make poor specimens.

Specimen preparation — At this point, you may decide to take the specimen to a Museum collection as is (in Kansas, the Sternberg Museum of Natural History or the University of Kansas Biodiversity Institute) or to prepare the specimen yourself. If you decide to prepare the specimen yourself, a Museum may provide you with the chemicals and equipment necessary, or at least assist you in obtaining them. Preparing specimens is time-consuming and Museums welcome the assistance.

• Tissue Collection — Tissue samples should be taken from every specimen. A sample of liver, heart, or skeletal muscle stored in a vial/tube of 95% (190 proof) ethanol labeled with the collector/museum field/catalog number. Tubes (source) with screw tops and a circular flange on the bottom, so they will stand upright, are ideal. The tissue extraction instruments should be run through a flame after each specimen to avoid cross-contamination. Avoid clipping toes or other external appendages as this reduces the number of external morphological characters available for study. Tissues collected and stored out of the light by this method have a long shelf-life.
• Tags — You will need to facilitate a means to associate your specimen with its collection data (date, locality, collector, etc). This could be done by placing each specimen in a separate jar and labeling it with the data. However, this most easily accomplished by affixing a small sequentially numbered tag (source) to each specimen with cotton string and keeping a separate catalog of the tag number and specimen data. Many field herpetologists get tags with a personal identifier and sequential number (e.g. initials [TWT 0827]). These field tags are permanently affixed to the specimen, and when the specimen is processed at the Museum, its tag will be added.
• Fixation — Formalin fixation is necessary to harden specimens before their transfer to their ultimate storage medium. Formalin (an aqueous solution of formaldehyde gas) can be obtained from most drug stores (source). Formalin is a known carcinogen and you should minimize your exposure to it. Rubber gloves are a necessity when working with formalin, as is operating in a well-ventilated area. Prepare formalin as a 10% solution. Specimens must be thawed or fresh. Prepare an appropriately sized Tupperware container with paper towels on the bottom. Attach a tag, or label the specimens in a manner so that the correct tissues and specimen can be associated later.
  • Reptiles — Inject formalin into the body cavity at regularly spaced intervals to ensure that the fixative comes in contact with internal tissues, but not so much as to distend the body. On males, inject formalin into the tail in an attempt to expose the hemipenes. Arrange the specimens in the container as follows, and cover them with more paper towels. The purpose of arranging the specimens is to make as many external characters as possible accessible. Spray/pour formalin over the towels to moisten them.
    • Snakes — Coil them in a spiral with the head on the outside.
    • Lizards — Spread out the arm, legs, fingers, and toes. If the specimen is greater than 6 inches in length, fold the tail alongside the body.
    • Turtles — Make sure the arms, legs, and head are outside of the shell. Place a block of wood into the mouth of the specimen so that it hardens with the mouth open.
  • Amphibians — Do not need to be injected. Arrange them in the container as follows for hardening.
    • Frogs — Make sure the arms, legs, fingers, and toes are spread apart.
    • Salamanders — Spread out the arm, legs, fingers, and toes. If the specimen is greater than 6 inches in length, fold the tail alongside the body.
    • Eggs — Place them in an appropriately sized container of 10% formalin directly.
  Specimens will require one to five days to harden. Check them periodically. When they are stiff, remove them from the tray and rinse them in a basin of cold water for 24 hours.
• Storage — Once rinsed, specimens should be transferred to jars containing 70% (150 proof) non-denatured ethanol (e.g Everclear) which can be purchased in most liquor stores... or from chemical supply companies in bulk. If the jars aren't transferred to a museum soon thereafter, it may be necessary to replace the ethanol periodically. This is especially true at first, as the ethanol will be diluted by the water associated with the rinsed specimens.